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From the Laboratory for Microvascular Research and Vascular Tissue Engineering, Institute of Reconstructive Plastic Surgery, New York University Medical Center, New York, New York
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Abstract |
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Impaired wound healing is a significant clinical problem, with an economic cost totaling billions of dollars yearly. One of the most common disease states associated with impaired tissue repair is diabetes mellitus. The mechanisms by which diabetes impedes tissue repair remain unclear. Previous studies have suggested decreased growth factor [keratinocyte growth factor, vascular endothelial growth factor (VEGF), platelet-derived growth factor] production, excess protease activity, or an increased microbial load as possible etiologies for the impaired wound healing observed in diabetes mellitus.4-6 In addition, diabetic patients often have co-existing peripheral vascular disease and polyneuropathy that further impede wound healing.
Although many studies have correlated imbalances or deficiencies in growth factor expression or protease activity with chronic wounds in vivo, few studies have examined the in vitro behavior of cells harvested from unwounded diabetics in processes integral to tissue repair. The evidence demonstrating that diabetes does have effects on essential aspects of fibroblast biology, such as proliferation and collagen synthesis,7-9 supports the need for further examination of more complex fibroblast functions during tissue repair. For example, fibroblasts produce matrix metalloproteinases (MMPs), which are critical for cell migration and tissue remodeling during wound healing. MMPs have been observed to be elevated in wounds of healing impaired mice and in wound fluid from chronic wounds.10-12 However, potential mechanisms for the increased production have not been specifically examined.
Fibroblasts also secrete numerous growth factors involved in the wound healing process, including VEGF.13-15 VEGF is absolutely essential for vascular development in utero, and its importance in tissue repair in the adult organism is highlighted by studies demonstrating deficiencies in tissue repair in models in which VEGF expression is experimentally reduced.16,17 The diminished neovascularization observed in animal models of impaired wound healing is in part thought to be the result of delayed or diminished production of VEGF and other angiogenic growth factors.18 However, the mechanism underlying this impairment is unclear.
To study the possible pathophysiological effect of the diabetic state on the effector cells of wound healing, we analyzed the behavior of early-passage fibroblasts harvested from the leptin receptor-deficient db/db mouse. This well-established model of type II diabetes is characterized by hyperglycemia, obesity, and well-documented impairments in wound repair.19-21 The db/db fibroblasts were compared to age- and passage-matched fibroblasts cultured from nondiabetic, wild-type mice of the same background strain. We specifically examined migration, MMP production, and the production of VEGF. Because wound healing is thought to occur in a relatively hypoxic environment22,23 we further analyzed these variables in both normoxic and hypoxic conditions. We demonstrate that fibroblasts from adult db/db mice maintain selective impairments in multiple cellular processes, and that these impediments to repair are accentuated by exposure to low oxygen tension. Notably, these deficits in VEGF production are acquired only after the onset of hyperglycemia and are therefore not directly attributable to the absence of the leptin receptor. These cellular abnormalities may play a role in the impaired wound healing observed in diabetics and may serve as future targets for intervention.
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Materials and Methods |
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Dulbecco’s modified Eagle’s medium, fetal bovine serum (FBS), trypsin, collagenase I, fibronectin, phosphate-buffered saline (PBS), Hanks’ balanced salt solution, and antibiotics for cell culture were all obtained from Life Technologies Inc. (Grand Island, NY). Type I rat-tail collagen was obtained from BD-Biosciences (San Jose, CA). Unless otherwise stated, all other chemicals were purchased from Sigma Chemical Co. (St. Louis, MO) and Fisher Scientific (Pittsburgh, PA).
Diabetic Animal Model
Eight- to 12-week-old female, pathogen-free, genetically diabetic C57BLKS/J-Leprdb (db/db) mice and their C57BLKS/J wild-type littermates were obtained from Jackson Laboratories (Bar Harbor, ME). Heterozygote breeder mice from the C57BLKS/J-Leprdb strain were also obtained from Jackson Laboratories and bred under standard conditions to obtain neonatal db/db and wild-type mice. Neonatal db/db homozygotes were identified by polymerase chain reaction genotyping as performed according to the protocol provided by the supplier (The Jackson Laboratory, Jackson, ME) (http://aretha. jax. org/pub-cgi/protocols/protocols. sh?objtype=protocol&protocol_id=348). The animals were housed in a central animal care facility maintained at 24°C, 30% humidity, and a 12-hour light-dark cycle. Water and standard rodent laboratory chow were available ad libitum. Body weights and nonfasting blood glucose levels were recorded at 2-week intervals from 2 weeks of age until 20 weeks. All procedures were approved by the Animal Care Committee of New York University.
Cell Culture
Primary isolates of dermal fibroblasts were harvested from separate neonatal and adult mice for each experiment. The animals were sacrificed and trunk skin was removed by sharp dissection. Special care was taken to remove the underlying adipose tissue. The harvested skin was then minced and digested for 2 hours in 0.20% collagenase I solution in serum-free Dulbecco’s modified Eagle’s medium at 37°C. The dissociated cells were then centrifuged and resuspended in a culture medium of Dulbecco’s modified Eagle’s medium with 10% FBS and 1% antibiotic/antimycotic supplement. The cells were grown at 37°C with 100% humidity in 5% CO2 in air. Medium was changed every other day; cells were passaged before they attained confluence. Experiments were performed using cultured cells at passage 3 or less and the experiments were repeated using cell lines isolated from different animals.
In Vitro Hypoxia
Hypoxic conditions were achieved via a custom-designed incubator with an attached hypoxic workspace (Reming BioInstruments, Redfield, NY). Hypoxia was achieved by the infusion of calibrated nitrogen gas into both the incubator as well as the attached workspace to achieve a constant oxygen concentration of 1%. The carbon dioxide concentration was maintained at 5%, and the air within the incubator was humidified and kept at 37°C. Before any experiment in the hypoxic incubator, media was placed within the incubator for 12 hours to pre-equilibrate the dissolved oxygen concentration within the media to 1% O2.
Migration Assays
Two independent migration assays were used to quantitate migration in response to extracellular matrix molecules. The first was a gold-salt phagokinetic migration assay, which was performed according to the method of Albrecht-Buehler,24 as modified by O’Keefe and colleagues.25 Briefly, glass coverslips were placed into an eight-well dish and coated with a colloidal gold salt solution. They were then coated with either: type I rat tail collagen ranging in concentration from 0 to 20 µg/ml, fibronectin ranging in concentration from 0 to 40 µg/ml, or laminin ranging in concentration from 0 to 60 µg/ml. Fibroblasts (n = 5000) were plated onto each coverslip in Dulbecco’s modified Eagle’s medium with 10% FBS and incubated at 37°C for 20 hours. The coverslips were rinsed with Hanks’ balanced salt solution and the cells were fixed. The cells were then visualized under dark-field optics. To quantitate migration, 10 high-power fields at x40 magnification were digitally photographed. The migration tracks of the cells were visible as black empty spaces against the background of bright gold-salt particles. The track area of the cells was calculated in pixels using Scion image analysis software.
The second assay was a modified Boyden chamber haptotaxis assay.26 Transwell plates (8-µm pore) (Costar; Corning, Corning, NY) were coated with either 10 µg/ml of type I collagen, 20 µg/ml of fibronectin, 30 µg/ml of laminin, or 0.5% bovine serum albumin solution. Cells (n = 75,000) were added to the upper chamber of each migration well and were allowed to migrate for 3 hours. After gentle removal of the nonmigratory cells from the filter surface of the upper chamber, the cells that migrated to the lower side were fixed, stained with crystal violet, and washed twice with PBS. The stain was then eluted with 10% acetic acid and the wavelength absorbance was measured at 600 nm on a Spectramax 340 plate reader. Background migration was controlled for by subtracting the absorbance value measured in the plates coated with 0.5% bovine serum albumin alone without extracellular matrix. Some experiments were conducted under hypoxic (1% O2) conditions.
Gelatin Zymography
Diabetic db/db and normal dermal fibroblasts in culture conditions described above were washed twice with PBS and the medium was changed to a serum-free fibroblast basal medium (Clonetics, San Diego, CA). The cells were serum-starved for 24 hours. Tissue culture dishes were coated with type I rat tail collagen at a concentration of 10 µg/ml. The fibroblasts were then plated onto these dishes at a density of 20,000 cells/cm2 and allowed to attach for 4 hours in fibroblast basal medium. After 4 hours the medium was removed and the cells were washed twice with PBS. The medium was then changed to a serum-free fibroblast growth medium (Clonetics) that was supplemented with 1 µg/ml of human fibroblast growth factor and 5 mg/ml of insulin. After 20 hours of incubation the conditioned medium was harvested and phenylmethyl sulfonyl fluoride and N-ethylmaleimide were added to a final concentration of 1 mmol/L and 2 mmol/L, respectively. The conditioned medium was then centrifuged for 1 hour at 5000 relative centrifugal force (RCF) and the protein content was normalized between samples. Gelatinolytic activity of conditioned medium from cultured cells was analyzed by electrophoresis in 10% sodium dodecyl sulfate-polyacrylamide gels containing 1 mg/ml of gelatin (Bio-Rad Ready Gels; Bio-Rad, Richmond, CA). After electrophoresis, the gels were soaked for 1 hour in 2.5% Triton X-100 at room temperature and then incubated overnight in digestion buffer (50 mmol/L Tris-HCl, pH 7.6, 0.2 mol/L NaCl, 5 mmol/L CaCl2, and 0.2% Brij-35). Subsequently the gels were stained using a single-step staining/destaining method.27 The bands representing specific activity of both MMP-2 and MMP-9 were then quantified via densitometry using Kodak 2D image analysis software (Eastman-Kodak, Rochester, NY).
MMP Enzyme-Linked Immunosorbent Assay (ELISA)
Conditioned medium from diabetic db/db and normal dermal fibroblasts in culture was harvested as described above. Total MMP-2 and MMP-9 concentration was determined using the commercially available Biotrak ELISA (Amersham Pharmacia, Piscataway, NJ). Standard curves were generated using serial dilutions of 100 ng/ml of stock standards supplied with the assays. The wavelength absorbance was measured at 405 nm on a Spectramax 340 plate reader and the total MMP concentration was determined by comparison to a standard curve.
VEGF ELISA
Production of VEGF by fibroblasts from newborn and adult, diabetic and normal, mice was measured using an ELISA technique for murine VEGF (Quantikine M Murine; R&D Systems, Minneapolis, MN). VEGF production by cells in culture was assayed in serum-free basal medium incubated in standard 21% O2 conditions (normoxia) or in 1% O2 (hypoxia). The hypoxic conditions were achieved as described above. Serum-free FBS that had previously equilibrated to the O2 tension of the respective experimental conditions was used throughout the entire experiment. Conditioned medium from diabetic db/db and normal dermal fibroblasts in culture was harvested as described above for zymography and protease inhibitors were added. The conditioned medium was then concentrated and the protein content was normalized. The VEGF concentration was determined using the Quantikine ELISA kit according to the manufacturer’s instructions.
Protein Assay
For protein quantitation the Coomassie Plus-200 Protein Assay Reagent (Pierce, Rockford, IL) was used. Protein level normalization for VEGF and MMP ELISA as well as for zymography was performed after concentration using Centricon-10 centrifugal filter devices (Amicon, Bedford, MA).
Proliferation Assay
To assess the proliferation rates of the diabetic and normal fibroblasts, the 5-bromo-2'-deoxy-uridine (BrdU) labeling and detection kit III (Roche, Indianapolis, IN) was used. The cells were plated at a density of 6400 cells/well into a 96-well plate and allowed to reach 70% confluence. BrdU was then added to the cells and incubated for 12 hours. A peroxidase labeled anti-BrdU antibody was then bound to the incorporated BrdU and the peroxidase substrate was then added. The catalysis of the peroxidase substrate by the enzyme yielded a colored reaction product. The wavelength absorbance of the samples was then measured at 405 nm on a Spectramax 340 plate reader and the absorbance of diabetic db/db fibroblasts was compared to that of normal fibroblasts.
Senescence Assay
ß-Galactosidase staining (Cell Signaling Tech, Beverly, MA) was used to quantify senescence at a pH of 6 where ß-galactosidase activity is only present in senescent cells.28,29 Briefly, the cells were first washed with PBS and then fixed at room temperature. They were then stained with a solution containing 5-bromo-4-chloro-3-indolyl-ß-D-galactopyranoside (X-gal) and allowed to incubate overnight at 37°C. The next day the cells were viewed under x200 magnification. Normal fibroblasts were compared to diabetic db/db fibroblasts for development of any blue color.
Statistical Analysis
All results are expressed as mean ± SEM of untransformed data. Controls for results expressed in percentile form were first averaged and then used for normalization. Statistical comparisons between groups were performed by Student’s t-test. P values <0.05 were considered significant. All experiments were repeated at least three times using cell lines isolated from at least three different mice of the same strain.
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Results |
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Differences in migration on type I collagen and fibronectin were observed between diabetic db/db and wild-type fibroblasts. Diabetic fibroblast migration was markedly impaired when compared to wild-type fibroblasts in both the gold-salt phagokinetic migration assay and the modified Boyden chamber haptotaxis assay. In the gold-salt phagokinetic migration assay (Figure 1)
wild-type fibroblast migration was greater than diabetic fibroblasts at all concentrations of type I collagen or fibronectin. At peak migration, wild-type fibroblast migration was fourfold greater (pixel area, 117,915.1 ± 10,284.7 versus 28,293.8 ± 3841.7; P < 0.001) than diabetic fibroblast migration (Figure 2)
. Similarly, when plated onto fibronectin, wild-type fibroblast migration was more than twofold (pixel area, 107,055.1 ± 6108.4 versus 51,033.9 ± 2871.5; P < 0.001) greater than diabetic fibroblasts (Figure 2)
. Neither cell type migrated to a significant degree when plated on laminin (data not shown).
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. Normal fibroblasts migrated 32% more (OD600, 0.469 versus 0.356; P < 0.05) than diabetic fibroblasts in response to fibronectin (Figure 3A)
. In hypoxic conditions (1% O2) wild-type fibroblast migration on type I collagen was up-regulated almost twofold (OD600, 0.211 versus 0.416; P < 0.05) whereas diabetic fibroblast migration was not significantly elevated (OD600, 0.119 versus 0.160; P = 0.1) (Figure 3B)
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MMP-9 activity was threefold greater (3.1 ± 0.2-fold, P < 0.001) in diabetic fibroblasts as assayed by gelatin zymography of the conditioned medium from cultured cells (Figure 4)
. This elevation seemed to be specific for the latent 92-kd form of MMP-9 because there was no measurable difference in gelatinolytic activity of the active 83-kd form of MMP-9. Conversely, there was no statistically significant difference in the gelatinase activity of MMP-2 (latent 72-kd or active 62-kd forms). These results were confirmed by ELISA (Figure 5A)
with significant elevation in MMP-9 concentrations demonstrated in conditioned medium from diabetic fibroblasts when compared to wild-type controls (0.816 ± 0.115 ng/ml versus 0.420 ± 0.024 ng/ml. P < 0.05). There was no difference detected by ELISA between diabetic and wild-type fibroblasts in production of total MMP-2 (Figure 5B)
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Differences between wild-type and diabetic fibroblasts were observed in the production of VEGF. Diabetic fibroblasts showed a severe impairment in VEGF production in all experimental conditions (Figure 6)
. Under standard oxygen tensions (21% O2) diabetic fibroblasts secreted less than 13% of the total VEGF produced by wild-type fibroblasts in basal media (4.5 ± 1.3 pg/ml versus 34.8 ± 3.3 pg/ml, P < 0.001). Under hypoxic (1% O2) conditions wild-type fibroblast production of VEGF was elevated threefold (101.9 ± 8.9 pg/ml versus 34.8 ± 3.3 pg/ml, P < 0.001). Diabetic fibroblasts, however, exhibited no significant up-regulation in VEGF production in response to the same hypoxic stimulus (11.8 ± 3.6 pg/ml versus 4.5 ± 1.3 pg/ml, P = 0.08).
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The proliferation of diabetic and wild-type fibroblasts was measured using BrdU incorporation. No significant differences were detected in the proliferation rate of diabetic and wild-type fibroblasts (OD450, 0.254 ± 0.022 versus 0.260 ± 0.024; P = 0.85). This assay measures the relative rate of DNA synthesis at a particular instant and is not averaged throughout time as with a growth curve. Therefore, it does not take into account growth variability that may be seen in culture throughout time.
Senescence was also examined and the results supported the proliferation data, revealing that the diabetic and wild-type fibroblasts were both nonsenescent in culture at the passages used. Less than 1% staining was detected in both cell types. No significant difference in senescence was noted (data not shown).
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Discussion |
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TopAbstractMaterials and MethodsResultsDiscussionReferences |
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Although it is well accepted that diabetic wounds heal poorly, the mechanisms underlying this phenomenon are not well understood. Descriptive observations of intact wounds have shed some light on possible mechanisms, but no single etiology has been proven to be responsible.32-34 Previous studies demonstrate that diabetic wounds have a deficiency of proangiogenic growth factors, a prolonged inflammatory state, and impaired cell migration and wound contraction. Several molecular mechanisms have been proposed to account for these disparate pathophysiological effects.35 Two in particular have relevance to the present study: the accumulation of advanced glycosylation end-products in cells exposed to chronic hyperglycemia, and oxidative damage resulting from overproduction of mitochondrial oxidative stressors.35 Both of these would likely result in permanent damage to cells, even when returned to a normoglycemic environment. As such, cells removed from chronically hyperglycemic animals should continue to exhibit abnormalities through early passages in vitro.
In this study, we have demonstrated that dermal fibroblasts harvested from diabetic mice exhibit abnormalities even when grown in an ex vivo culture environment that has been optimized for nutrient, growth factor, and glucose concentrations. We have demonstrated using two different in vitro assays of cell migration that diabetic fibroblasts do not migrate as well as wild-type fibroblasts. Because hypoxia is a known stimulus for increased migration in keratinocytes and fibroblasts,36,37 we repeated these assays under hypoxic conditions. We demonstrated that hypoxia is unable to increase diabetic fibroblast migration, implying that the hypoxic response may be impaired in these cells.
We examined gelatinase activity as a possible mechanism for the decreased migration in the db/db fibroblasts. We were surprised to find only a selective increase in pro-MMP-9 in db/db fibroblasts, with no difference in MMP-2 levels or active MMP-9. This seems unlikely to be responsible for the observed impairments in migration. However, MMPs have physiological roles in tissue repair other than cell migration and, indeed, pathologically elevated levels seem to impair normal wound healing. Evidence for this comes from previous studies that have demonstrated increased MMP-9 in chronic wounds,11,38 which is felt to produce the prolonged inflammatory state present in diabetic animal wound-healing models.39,40 Previously, it has been proposed that this excess proteolytic enzyme production is secondary to an increased microbial burden in vivo. However, we have demonstrated that elevated MMP-9 production by db/db fibroblasts can occur in the absence of infection. This pattern of increased pro-MMP-9 expression in settings of elevated glucose levels has also been described in bovine aortic endothelial cells,41 suggesting that elevated glucose levels specifically enhance MMP-9 secretion in certain cell types. The mechanism underlying the increased level of pro-MMP-9 levels remains unclear and is an area of ongoing investigation.
The most striking result in the present study is the markedly diminished production of VEGF in diabetic fibroblasts. Previous studies have described diminished levels of VEGF in vivo during wound healing in the db/db mouse,18 and the present study suggests that an etiology for the diminished VEGF level observed is a blunted fibroblast release of VEGF in response to injury. Further support for this hypothesis is that db/db fibroblasts are unable to up-regulate VEGF production in hypoxic (1% O2) conditions, whereas wild-type fibroblasts exhibited a threefold increase in VEGF production. To determine whether this impaired VEGF response was because of the diabetic state or resulted from the absence of the leptin receptor, we examined dermal fibroblasts harvested from neonatal db/db mice (before the onset of hyperglycemia), and compared these to their wild-type littermates. This was necessary because leptin has been implicated in processes including tissue repair and angiogenesis.42-48 Our results demonstrated no abnormalities or difference in VEGF production or response to hypoxia in db/db mouse fibroblasts obtained before the development of diabetes. We conclude that the differences observed in the adult db/db fibroblasts are because of development of the diabetic phenotype of these animals (hyperglycemia, insulin resistance, and obesity), occurring between 4 and 8 weeks of life. Clearly, the significant abnormalities in VEGF production at baseline and in response to hypoxia cannot solely be attributed to aberrant leptin signaling.
The abnormal VEGF activity and response to hypoxia may have a profound effect on the healing process in diabetic wounds, particularly because many diabetic wounds are located in ischemic extremities. Without an appropriate angiogenic response, the subsequent phases of cell proliferation and matrix deposition are delayed because the neovasculature in healing wounds is required for the deposition of matrix.14,49 This suggests that it may be possible to enhance wound healing in diabetic patients by reversing these discrete deficits, most obviously by supplementing them with exogenous VEGF. It is also interesting to consider whether diminished VEGF production may underlie the decreased tolerance of diabetic patients to ischemic events in a variety of other tissues, including heart, brain, and muscle.50-54
In conclusion, we have demonstrated selective impairments in aspects of fibroblast behavior important for tissue repair in fibroblasts harvested from diabetic db/db mice. These include impaired cell migration and MMP-9 overexpression and an inability of diabetic fibroblasts to produce normal levels of VEGF at baseline or in response to hypoxia. This suggests that cells are persistently altered in response to prolonged hyperglycemia, at least with respect to VEGF production. Studies are underway to elucidate how glucose and hypoxia interact to diminish angiogenic growth factor expression in fibroblasts and other cell types.
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Footnotes |
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Funded by a grant from the Sarnoff Endowment for Cardiovascular Research, Inc.
O.Z. Lerman and R.D. Galiano contributed equally to this work.
Accepted for publication October 3, 2002.
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